Mass spectrometry (MS) is a powerful analytical technique that is used to identify unknown compounds, to relatively quantify known compounds, and to elucidate the structure and chemical properties of molecules.
Traditionally, proteome analysis has been performed using a combination of high resolution gel electrophoresis, in particular, two-dimensional (2D) gel electrophoresis, to separate proteins, and mass spectrometry to identify proteins. Typically, separation is by isoelectric focusing, which separates proteins by charge in a pH gradient, followed by SDS-PAGE, which separates proteins by molecular weight. After staining and separation, the mixture appears as a two-dimensional array of spots of separated proteins. Spots are excised from the gel, enzymatically digested, and subjected to mass spectrometry for identification. Relative quantitation of the identified proteins can be performed by observing the relative intensities of the spots via image analysis of the stained gel. However, because ionization efficiency of different protein fragments varies greatly, comparative quantification using mass spectrometry is unreliable.
Alternatively, peptides have been labeled isotopically before gel separation and expression levels quantified by mass spectrometry or radiographic methods. Absolute concentrations have not been achievable using these methods.
2D gels have a number of drawbacks. In particular, the approach is sequential and tedious, and is additionally fundamentally limited in that many biologically important classes of proteins, such as nuclear proteins or membrane proteins, are practically undetectable using these methods. Very acidic or basic proteins, very large or small proteins, and membrane proteins are either excluded or underrepresented in 2D gel patterns. Low abundance proteins, including regulatory proteins, are rarely detected when entire cell lysates are analyzed, reflecting a limited dynamic range. These deficiencies are detrimental for quantitative proteomics.
Protein standards for 2D gels are generally known. 2D gels separate proteins and polypeptides based on size. For example, US Patent application 20030157720 discloses a protein standard set that is based upon commercial protein standards providing polypeptides as standards for calculating molecular weight and the amount of peptide present.
However, in mass spectrometry, separation is not based on size alone, but separation is based on a mass to charge ratio. The mass to charge ratio is accurately obtained by MS. However, MS fragments are not easily assigned to a particular protein without sequencing (which is possible with many popular MS machines). Different peptides have different ionization efficiencies and patterns for MS, which prevents accurate comparison of one protein or polypeptide to another as is seen with 2D gels. An ideal MS standard should behave identically to the protein being measured during preparation for MS as well as within the MS.
Because it can provide detailed structural information, mass spectrometry is currently a valued analytical tool for biochemical mixture analysis and protein identification. For example, capillary liquid chromatography combined with electrospray ionization tandem mass spectrometry has been used for large-scale protein identification without gel electrophoresis. Qualitative differences between spectra can be identified, and proteins corresponding to peaks occurring in only some of the spectra are considered as candidate biological markers. Mass spectrometry analyses are not quantitative, however. In most cases, quantitation in mass spectrometry requires an internal standard, a compound introduced into a sample at known concentration. Spectral peaks corresponding to sample components are compared with the internal standard peak height or area for quantitation. Ideally an internal standard has elution and ionization characteristics similar to and preferably identical to those of the target compound but preferably the standard generates ions with a detectably different mass-to-charge ratio.
Using internal standards for complex biological mixtures is problematic. In many cases, the compounds of interest are unknown a priori, preventing appropriate internal standards from being devised. The problem is more difficult when there are many compounds of interest. In addition, biological samples are often available in very low volumes, and addition of an internal standard can dilute mixture components significantly. Low-abundance components, often the most relevant or significant ones, may be diluted to below noise levels and hence undetectable. Also, it can be difficult to judge the proper amount of internal standard to use. Thus internal standards are not widespread solutions to the problem of protein quantitation.
To reliably identify a biomolecule, such as a protein, from a sample, mass spectrometry (MS) based methods for proteomics rely on identification of a fragment ion, for example a peptide generated by the sequence specific fragmentation of a protein. Therefore, biomolecules, such as proteins and carbohydrates generally need to be enzymatically or chemically fragmented prior to mass spectrometric analysis. A protein generally generates a large number of peptides and hence a large number of peptides must be sequenced for each experiment.
Stable isotope labeling with amino acids in cell culture (SILAC) was developed as a useful tool for assaying relative concentrations of proteins of cells grown in culture. SILAC incorporates a label into proteins for mass spectrometric (MS)-based proteomics. SILAC relies on metabolic incorporation of a “light” or “heavy” form of an amino acid into proteins.
In a SILAC experiment, two groups of cells are grown in culture media that are essentially identical except in one respect: one media contains a “light” and the other a “heavy” form of a particular amino acid (for e.g. L-leucine or deuterated L-leucine). Thus, conventional SILAC techniques rely on growing parallel cultures where one set of cultures is grown in media containing an isotopically-labeled amino acid (such as 15N-Arg) and the other culture set is grown in conventional media thereby allowing an investigator to challenge one set of cultures with an external stimulus to monitor the relative changes in expression. With each cell doubling the cell population replaces at least half of the original form of the amino acid, eventually incorporating 100% of a given “light” or “heavy” form of the amino acid. Thus, when the labeled analog of an amino acid is supplied to cells in culture instead of the natural amino acid, it is incorporated into all newly synthesized proteins. After a number of cell divisions, each instance of this particular amino acid will be replaced by its isotope-labeled analog. Because there little chemical difference between the labeled amino acid and the natural amino acid isotopes, the cells behave like the control cell population grown in the presence of normal amino acid.
Conventional SILAC processes two culture lysates that are mixed and proteolyzed with optionally one or more stages of affinity enrichment. Unlabeled and labeled samples can be combined prior to lysis of the cells and treated as a single sample in all subsequent steps. This allows the experimenter to use any method of protein or even peptide purification (after enzymatic digestion) without introducing error into the final quantitative analysis.
SILAC methods are disclosed for example, in U.S. Pat. No. 6,391,649 to Chait. In SILAC, quantitation by mass spectrometry (MS) is performed by measuring the relative peak intensities of the heavy-labeled and the light-labeled isoforms of the peptides. Unlike chemical labeling techniques (e.g. isotopically coded affinity tagging (ICAT) or iTRAQ, Applied Biosystems, Framingham Mass.), the incorporation of isotopically heavy amino acid is nearly 100%. The difference in the mass of the isotope in each cell pool results in two distinct, closely spaced peaks for each protein or peptide actively produced by the samples in the mass spectrum. One peak corresponds to a protein or peptide from a protein from the cell pool with the normal abundance of isotopes. The other peak corresponds to a protein or peptide from the cell pool enriched in one or more of the isotopes. A ratio is computed between the peak intensities of at least one pair of peaks in the mass spectrum. The relative abundance of the protein in each sample may be determined based on the computed ratio. The protein may be identified by the mass-to-charge ratios of the peaks in the mass spectrum, as well as by other means known in the art.
Up to the point of the MS, none of the steps of the Chait process discriminates between a protein that contains the natural abundance of isotopes from the same protein from the enriched sample. Thus, the ratios of the original amounts of proteins from the two samples are maintained, normalizing for differences between extraction and separation of the proteins in the samples.
With labeled cells in SILAC, one can proceed to do sub-cellular purification of intact organelle structures or multi-protein complexes. The two samples can be combined as whole cells and a single subcellular preparation of nuclei, mitochondria, etc., then prepares the samples (now combined) for MS. Any sample preparation bias introduced by the comparison of two separate preparation steps (as would be the case in a chemical modification method) is avoided. SILAC is thus proven as a useful tool for proteomic analysis.
An example of a typical SILAC experiment is illustrated in FIG. 1. FIG. 1 depicts two identical cultures grown in parallel where one culture is grown under media conditions that enrich the 15N isotope of an amino acid (such as arginine) and the other is grown in conventional media. One of the cultures is challenged by an external stimulus (such as application of a drug). After the stimulus, the cell cultures are processed and lysates are produced. The lysates from each culture are mixed and proteolyzed with an enzyme such as trypsin. After some chromatography, the sample is analyzed by MS. MS analysis is capable of resolving the sample components by mass and measuring the relative abundance of the light and heavy isoforms of a specific peptide. The light and heavy isoforms will appear as two peaks differing in mass by the difference in mass due to the heavy isotope present. Proteins unaffected by the challenge will have a like ratio of the heavy isoform to light isoform peaks. A different ratio of the heavy isoform to light isoform indicates which culture (challenged or control) has increased or decreased expression of that protein.
Despite the advantages of SILAC over conventional chemical labeling techniques for quantifying expressed proteins, there are several limitations of the technique. As the name of the technique implies, incorporation of the heavy isotope requires protein expression in active cell cultures. Thus, the technique cannot be applied to tissue samples, biopsies or tissue slices. Another limitation is that protein quantitation by SILAC is merely relative (i.e. the relative expression of a specific protein in one experimental condition versus another) in like treated parallel samples. SILAC does not provide quantitation information to allow comparison of concentration of different molecules, for example different proteins. Absolute concentration information is not obtained using SILAC. Further, because SILAC does not provide absolute quantitation, a comparison of results between experiments is difficult to analyze.
Accordingly, MS and SILAC are useful tools. However, MS analysis does not lend to easy absolute quantitation, but only relative quantitation of the molecules analyzed. SILAC suffers from its ability to only provide information of relative concentration comparing protein concentration between samples, and absolute quantity information may not be obtained using SILAC.